Patients with tuberculosis disease have Mycobacterium tuberculosis-specific CD8 T cells with a pro-apoptotic phenotype and impaired proliferative capacity, which is not restored following treatment

Abstract

CD8 T cells play a critical role in control of chronic viral infections; however, the role of these cells in containing persistent bacterial infections, such as those caused by Mycobacterium tuberculosis (Mtb), is less clear. We assessed the phenotype and functional capacity of CD8 T cells specific for the immunodominant Mtb antigens CFP-10 and ESAT-6, in patients with pulmonary tuberculosis (TB) disease, before and after treatment, and in healthy persons with latent Mtb infection (LTBI). In patients with TB disease, CFP-10/ESAT-6-specific IFN-γ+ CD8 T cells had an activated, pro-apoptotic phenotype, with lower Bcl-2 and CD127 expression, and higher Ki67, CD57, and CD95 expression, than in LTBI. When CFP-10/ESAT-6-specific IFN-γ+ CD8 T cells were detectable, expression of distinct combinations of these markers was highly sensitive and specific for differentiating TB disease from LTBI. Successful treatment of disease resulted in changes of these markers, but not in restoration of CFP-10/ESAT-6-specific CD8 or CD4 memory T cell proliferative capacity. These data suggest that high mycobacterial load in active TB disease is associated with activated, short-lived CFP-10/ESAT-6-specific CD8 T cells with impaired functional capacity that is not restored following treatment. By contrast, LTBI is associated with preservation of long-lived CFP-10/ESAT-6-specific memory CD8 T cells that maintain high Bcl-2 expression and which may readily proliferate.

Figure 1. CFP-10/ESAT-6-specific CD8 T cells in patients with TB have a short-lived, pro-apoptotic phenotype.

PBMCs from individuals with LTBI (n = 18) and patients with TB disease (n = 20) were stimulated for 6 hours with CFP-10 and ESAT-6 peptide pools; intracellular IFN-γ production was measured in CD8 T cells by flow cytometry. (A) Frequencies of CFP-10 and ESAT-6-specific IFN-γ⁺ CD8 T cells in persons with LTBI and patients with TB disease. Horizontal lines represent the median. Data are shown after subtraction of background cytokine production in the negative control condition. The dotted line separates individuals who met the criteria for a positive CD8 T cell response for further phenotypic analyses (above dotted line: n = 6 LTBI; n = 13 TB disease), and individuals in whom the frequency of IFN-γ⁺ CD8 T cells was too low to meet the criteria for further phenotypic analyses (below dotted line). (B) Representative flow cytometry data from an individual with LTBI and a patient with TB disease, following stimulation of PBMCs with an ESAT-6 peptide pool. Grey cells indicate the total CD8 T cell population (gated on VIVID^l°CD3⁺CD8⁺ cells); black cells indicate ESAT-6-specific CD8 T cells (gated on VIVID^l°CD3⁺CD8⁺IFN-γ⁺ cells). (C) Summary data of the percentage of specific IFN-γ⁺ CD8 T cells expressing Ki67, Bcl-2, CD127, CD57 and CD95 in individuals with LTBI and TB disease. Differences were assessed using the Mann-Whitney test. § indicates p values that did not remain significant after applying the Bonferroni correction for multiple comparisons. (D) Comparison of the expression of intracellular Ki67 and Bcl-2 between the total CD8 T cell population and specific IFN-γ⁺ CD8 T cells within the same individual. Only individuals meeting the criteria for a positive CD8 T cell response in the ICS assay were included in this paired analysis (n = 6 LTBI; n = 13 TB disease). Differences were assessed using the Wilcoxon matched-pairs test.

Figure 2. Differential co-expression patterns of Bcl-2, CD57 and CD95 by CFP-10/ESAT-6-specific CD8 T cells distinguish individuals with LTBI and patients with TB.

Boolean analysis was performed to determine co-expression patterns of Bcl-2, CD57, and CD95 by CFP-10 and ESAT-6-specific CD8 T cells detected by ICS, as described in Figure 1. (A) Co-expression patterns of Bcl-2, CD57, and CD95 by specific IFN-γ⁺ CD8 T cells detectable in individuals with LTBI (open bars; n = 6) and patients with TB disease (grey bars; n = 13). Cells were gated on VIVID^l°CD3⁺CD8⁺IFN-γ⁺ cells. Differences were assessed using the Mann-Whitney test. § indicates p values that did not retain statistical significance after applying the Bonferroni correction for multiple comparisons. Pie graphs indicate the median proportion of each population contributing to the total specific IFN-γ⁺ CD8 T cell response. (B) Comparison of the proportion of Bcl-2⁺CD57⁻CD95⁺ cells contributing to the total CFP-10 and ESAT-6-specific CD8 T cell responses in individuals with LTBI and patients with TB disease. Horizontal lines represent the median. Differences were assessed using the Mann-Whitney test. The dotted line indicates the cut-off (18.7%) that distinguishes individuals with LTBI and TB disease, with 100% specificity and 100% sensitivity. (C) Receiver operator characteristic (ROC) curve indicating the sensitivity and specificity of the proportion of CFP-10/ESAT-6-specific CD8 T cells that are Bcl-2⁺CD57⁻CD95⁺ to distinguish individuals with LTBI and TB disease. An area under the ROC curve (AUC) analysis was performed to further evaluate the performance of this phenotypic expression profile in distinguishing individuals with LTBI and TB disease.

Figure 3. Pro-apoptotic phenotype of CFP-10/ESAT-6-specific CD4 T cells in TB disease is characterized by increased expression of Ki67 and CD95 and downregulation of Bcl-2.

Expression of Ki67, Bcl-2, CD127, CD57, and CD95 by specific CD3⁺CD8⁻IFN-γ⁺ T cells was measured by flow cytometry, as described in Figure 1. Greater than 90% of CD3⁺CD8⁻IFN-γ⁺ T cells following stimulation with CFP-10 and ESAT-6 peptide pools were found to be CD4⁺ (see Fig. S3), and are referred to here as CD4 T cells. Phenotypic analysis was performed on individuals with positive CD4 T cell responses (defined as CD3⁺CD8⁻IFN-γ⁺) to either CFP-10 or ESAT-6 peptide pools (n = 16/18 with LTBI; n = 10/20 with TB disease). (A) Summary data of the percentage of specific IFN-γ⁺ CD4 T cells expressing Ki67, Bcl-2, CD127, CD57 and CD95 in individuals with LTBI and patients with TB disease. Differences were assessed using the Mann-Whitney test, followed by the Bonferroni correction for multiple comparisons. (B) Flow cytometry data indicating expression of Bcl-2 and Ki67 by ESAT-6-specific CD4 T cells in an individual with LTBI (top plot) and an individual with TB disease (bottom plot). Plots are shown gated on CD3⁺CD8⁻IFN-γ⁺ T cells. (C) Co-expression patterns of Bcl-2, CD95, and Ki67 by IFN-γ⁺ CD4 T cells in individuals with LTBI (open bars; n = 16) and TB disease (grey bars; n = 10). Differences were assessed using the Mann-Whitney test, followed by the Bonferroni correction for multiple comparisons. (D) Comparison of the proportion of Bcl-2⁺CD95⁻Ki67⁻ cells contributing to the total CFP-10/ESAT-6-specific CD4 T cell responses in individuals with LTBI and patients with TB disease. Horizontal lines represent the median. Differences were assessed using the Mann-Whitney test. The dotted line indicates the cut-off (7%) that distinguishes individuals with LTBI and TB disease, with 90% sensitivity and 75% specificity. (E) Receiver operator characteristic (ROC) curve indicating the sensitivity and specificity of the proportion of CFP-10/ESAT-6-specific CD4 T cells that are Bcl-2⁺CD95⁻Ki67⁻ to distinguish individuals with LTBI and TB disease. An area under the ROC curve (AUC) analysis was performed to further evaluate the performance of this phenotypic expression profile in distinguishing individuals with LTBI and TB disease. (F) Co-expression patterns of Bcl-2, CD95, and Ki67 by CMV pp65-specific CD4 T cells in individuals with LTBI (open bars) and TB disease (grey bars). Data are shown from 16 individuals with LTBI and 13 patients with TB disease who had positive CD4 T cell responses to a CMV pp65 peptide pool. No differences were found between individuals with LTBI and TB for any of the CMV-specific CD4 T cell subsets (Mann-Whitney test).

Figure 4. CFP-10/ESAT-6-specific CD8 T cell phenotype is associated with mycobacterial antigen load.

Phenotypic analysis of CFP-10 and ESAT-6-specific IFN-γ⁺ CD8 T cells was performed as described in Figure 1. Cells were analyzed prior to initiating anti-TB treatment (Pre-TB Tx), and 2 months and 6 months after initiation of treatment. (A) Flow cytometry data indicating Ki67 and Bcl-2 expression by IFN-γ⁺ ESAT-6-specific CD8 T cells from a TB diseased patient at two time points: prior to treatment and 6 months after initiation of treatment, corresponding to the end of the treatment period. (B) Representative histogram overlays indicating expression of Bcl-2, CD127 and CD95 within CFP-10/ESAT-6-specific IFN-γ⁺ CD8 T cells. Black lines indicate expression prior to treatment; grey lines indicate expression 6 months after initiation of treatment. The numbers in the upper right corner of the histograms indicate the median fluorescence intensity (MFI) at each time point (black: pre-treatment MFI; grey: 6-month TB treatment MFI). Data shown in panels A and B are gated on VIVID^l°CD3⁺CD8⁺IFN-γ⁺ cells. (C) Summary data comparing the expression of Ki67, Bcl-2, CD127 and CD95 on CFP-10/ESAT-6-specific IFN-γ⁺ CD8 T cells from the same individual at two time points: prior to treatment (Pre-TB Tx) and at the end of the 6-month treatment period (6 mo. TB Tx). Data are shown from 7 individuals who maintained positive CFP-10/ESAT-6-specific CD8 T cell responses throughout the 6-month duration of treatment. Differences between time points were assessed using the Wilcoxon matched pairs test. § indicates p values that did not retain statistical significance after applying the Bonferroni correction for multiple comparisons. The proportion of CFP-10/ESAT-6-specific IFN-γ⁺ CD8 T cells that express Ki67 (D), CD95 (E), Bcl-2 (F), and CD127 (G) are shown for individuals with LTBI (n = 6), and TB diseased patients prior to treatment (n = 13; Pre-TB Tx), and 2 months (n = 10; 2 mo. TB Tx) and 6 months (n = 8; 6 mo. TB Tx) following initiation of treatment. Differences in panels D-G were first assessed using a Kruskal-Wallis test, followed by a Dunn's post-test to correct for multiple comparisons; § indicates p values that did not remain significant following correction with the Dunn's post-test.

Figure 5. Lack of restoration of CFP-10/ESAT-6-specific CD4 and CD8 T cell proliferative capacity following completion of treatment.

The proliferative capacity of CFP-10/ESAT-6-specific CD4 and CD8 T cells was measured following a 6-day stimulation of freshly isolated PBMCs with CFP-10 and ESAT-6 peptide pools. (A) Flow cytometry data of the proliferative capacity of antigen-specific CD8 T cells from a TB diseased patient analyzed at 4 time points: prior to treatment (Pre-TB Tx), 2 months of treatment (2 mo. TB Tx), 6 months of treatment (6 mo. TB Tx, corresponding to the end of the treatment period), and 6 months after completion of treatment (12 mo. TB Tx). The cells shown are gated on VIVID^l°CD3⁺CD8⁺ lymphocytes. The percentage on each plot indicates the percentage of proliferating CD8 T cells after subtraction of background proliferation in the negative control condition. (B, D) Longitudinal analysis of the proliferative capacity of CFP-10 and ESAT-6-specific CD8 (panel B) and CD4 (panel D) T cells in a subset of TB diseased patients who were followed for up to one year after TB diagnosis (n = 19 with at least 3 time points available for analysis). There were no significant differences in the frequency of proliferating antigen-specific CD8 or CD4 T cells over time (Kruskal-Wallis test). (C, E) Cross-sectional comparison of the CFP-10 and ESAT-6-specific CD8 (panel C) and CD4 (panel E) T cell proliferative capacity between individuals with LTBI (n = 34) and active TB disease at 4 time points (pre-TB tx: n = 34; 2 months of TB tx: n = 19; 6 months of TB Tx [corresponding to the end of the treatment period]: n = 25; 12 months post initiation of TB tx [corresponding to 6 months after the end of the treatment period]: n = 10). Data are shown after subtraction of background proliferation in the negative control condition. Differences in panels C and E were first assessed using a Kruskal-Wallis test, followed by a Dunn's post-test to correct for multiple comparisons. All comparisons remained significant following correction with the Dunn's post-test.